West Nile virus (WNV) was introduced to North America two decades ago, but for many species, including Ruffed Grouse (Bonasa umbellus), the effects of WNV on individuals and populations remain poorly understood. Recent studies suggest the effect of WNV on Ruffed Grouse might vary among geographic regions, depending on habitat conditions. We studied WNV in Minnesota, US, during 2018–19, in a region known to have abundant Ruffed Grouse habitat and a population cycling around a stable long-term average. We worked with cooperating hunters to collect hearts, feathers, and blood on filter strips from birds harvested in the fall to examine exposure to the virus. We detected antibodies to WNV or a flavivirus (probably WNV) in 12.5% and 12.3% of birds in 2018 and 2019, respectively. However, we did not isolate the virus from any heart samples, indicating that exposed birds were not experiencing an active infection of WNV at the time of harvest. Our findings indicate that, although Minnesota Ruffed Grouse are exposed to WNV, some birds mount a successful immune response and survive. However, our sampling approach did not account for birds that might have become infected over the summer and died, so it is unknown how much WNV mortality occurred before the fall harvest. Birds lost to WNV over the summer could reduce the number of birds that hunters see in the fall, thus reducing the quality of their hunting experiences. Management options for mitigating WNV impacts and other stressors consist primarily of providing high-quality Ruffed Grouse habitat that produces birds in good condition that are more likely to recover from infection.

West Nile virus (WNV), an arthropod-borne virus of the Flavivirus genus, originated in the eastern hemisphere and was first detected in the US in 1999 (Eidson et al. 2001). It is now established throughout the lower 48 states, with reports in >300 bird species (CDC 2017; Ahlers and Goodman 2018). Some avian species readily die of WNV infection, such as American Crows (Corvus brachyrhynchos), Blue Jays (Cyanocitta cristata), House Sparrows (Passer domesticus), and Common Grackles (Quiscalus quiscula); however, infected birds of most species can survive (Komar et al. 2003). In some birds, mortality events in the wild from WNV have never been documented, for example American Robins (Turdus migratorius) and chickadees (Poecile spp.; LaDeau et al. 2007; George et al. 2015). Other species, such as Field Sparrows (Spizella pusilla) and Red-eyed Vireos (Vireo olivaceus) initially had lower survival but then populations recovered.

The virus is transmitted among birds primarily by mosquitoes of the Culicinae family (Ahlers and Goodman 2018). Culex spp. of mosquitoes primarily feed on birds, but some species also feed on mammals, providing a bridge for WNV transmission from birds, which rapidly develop viremia, to humans and horses, which serve as dead-end hosts unable to pass the virus to mosquitoes (Molaei et al. 2006; Reisen 2012; Ahlers and Goodman 2018). The virus has been detected in as many as 66 mosquito species (CDC 2019) with different species acting as vectors in different hosts, geographies, seasons, and diurnal periods (Ahlers and Goodman 2018). The geographic range of some Culex spp. of mosquitoes is expanding with climate change, and the WNV transmission season is predicted to become longer (Hongoh et al. 2012; Chen et al. 2013; Morin and Comrie 2013), elevating concern for potentially vulnerable bird populations.

Recent concern about WNV effects on Ruffed Grouse (Bonasa umbellus), a popular forest gamebird, arose among hunters in Minnesota, US, when WNV was reported to be negatively related to population recovery in areas with poor quality habitat in Pennsylvania, US (Stauffer et al. 2018). In Minnesota, WNV was first documented in 2002 (Minnesota Department of Health), and in Ruffed Grouse in 2005 (Ruffed Grouse Society unpubl. data), but WNV-surveillance in grouse was negligible before 2018. Interest in WNV surveillance was instigated by news of the Pennsylvania study and a 7.8% decline in grouse harvest per active hunter during the 2017 Minnesota hunting season (Dexter 2018), in spite of a 57% increase in drums per stop, an index of relative population size, in the annual Ruffed Grouse drumming survey that had been conducted that spring (Roy 2017).

The Ruffed Grouse drumming survey has historically been reasonably predictive of the fall hunting season, with better-than-average hunting seasons occurring when the 10-yr cycle is near its peak as indicated by the index “drums per stop” (Ammann and Ryel 1963; Stoll 1980). However, this survey is conducted in the spring, before juveniles recruit to the fall population. A strong breeding population is usually related to strong juvenile production, but in years of poor chick survival, hunters may see and harvest fewer birds, because juveniles comprise most of the harvest (Dorney and Kabat 1960; Dorney 1963). Although the drumming survey indicates that the population has been cycling around a stable population average in the core of Minnesota Ruffed Grouse range (Roy 2017), some hunters have reported seeing fewer birds during recent peaks in the cycle, and this is reflected in state harvest data (Davros and Dexter 2019), leading to concern about chick survival. Previous studies have reported greater susceptibility of juvenile birds to WNV disease (Austin et al. 2004; Sovada et al. 2008; Cox et al. 2015) and juvenile birds having longer or higher-titered viremia than adults have (Nemeth and Bowen 2007; reviewed in Pérez-Ramírez et al. 2014). A discrepancy between hunter expectations and fall bird numbers could occur if WNV affects summer survival of young grouse, but mortality does not carry over to affect the breeding population the following spring because its effect is below the annual survival threshold set by other sources of natural mortality. This is much like the fact that hunting mortality does not affect the breeding population the following spring if harvest mortality is below the threshold set by natural mortality.

We expected that Ruffed Grouse in Minnesota would be exposed to WNV, based on annual reports of WNV in the state (Minnesota Department of Health 2020). Captive studies involving experimental WNV infection of Ruffed Grouse and Greater Sage-grouse (Centrocercus urophasianus) indicated that grouse are susceptible to disease (Clark et al. 2006; Nemeth et al. 2017) and that WNV could adversely affect chick survival in Ruffed Grouse (Nemeth et al. 2017). However, results obtained in captivity with small numbers of wild birds confined in artificial conditions do not necessarily reflect those in wild settings (Komar et al. 2003) and may overestimate morbidity and mortality in free-ranging birds in which exposure doses, repeated exposures, diet, social interactions, activity levels, stressors, and other variables may influence health outcomes (Pérez-Ramírez et al. 2014). For example, survival of Greater Sage-grouse infected with WNV has been documented in wild populations (Walker et al. 2007), despite 100% mortality in experimentally infected, non-vaccinated birds at <6 d of infection (Clark et al. 2006). Stress of confinement and repeated handling can suppress immune responses and contribute to mortality in wild animals in laboratory settings (Owen et al. 2012; Pérez-Ramírez et al. 2014). Moreover, Ruffed Grouse habitat is not limiting populations in Minnesota (Zimmerman et al. 2009; Kouffeld et al. 2013; Rusch et al. 2020), and abundant habitat might produce birds more resilient to severe WNV infection. Northern Minnesota is predominantly forested and much of that is aspen, in which Ruffed Grouse reach their highest densities (Gullion and Alm 1983; Rusch et al. 2020). In Pennsylvania, Ruffed Grouse persisted in areas with increasing habitat and low prevalence of WNV in mosquitoes, but had lower persistence in areas in which appropriate habitat was scarce and WNV prevalence was highest (Stauffer et al. 2018).

We partnered with Ruffed Grouse hunters to obtain samples from harvested birds and examine WNV exposure and infection in Minnesota. We estimated serologic exposure (seroprevalence) to WNV in juveniles and adults and tested hearts to determine prevalence of active (i.e., recent) infections.

Field data collection

In 2018, we focused sample collection within 96.6-km radii, centered on the three small cities of Grand Rapids, Longville, and Bemidji, Minnesota, to simplify logistics of sampling kit distribution to hunters. We also worked with organizers of special hunting events within those areas to collect samples. In 2019, we expanded sample collection to the entire forested region of Minnesota so that more hunters could participate and collect a representative sample. We made sampling kits available at Minnesota Department of Natural Resources (MNDNR) wildlife offices and during organized hunting events. We issued informational press releases, gave presentations to target audiences, and advertised the study in the 2018 and 2019 Minnesota Hunting Regulations and on the MNDNR website. The Ruffed Grouse Society and Pineridge Grouse Camp also offered raffle incentives to encourage voluntary hunter participation.

We provided hunters with sampling kits, including a protocol, a single Nobuto filter strip (Advantec®, Dublin, California, USA) for blood collection, a Whirl-Pak (Madison, Wisconsin, USA) for heart storage, a Ziploc bag (Racine, Wisconsin, USA) for feathers, and a datasheet. We asked hunters to collect hearts in a manner similar to that of previous WNV studies in wild birds (Komar et al. 2003; Nemeth et al. 2007) and for ease of identification and collection. The protocol included directions and images to determine the sex and age of birds based on feather characteristics (DeStefano et al. 2014) and to collect blood on filter strips (Nemeth et al. 2021). We requested information on harvest date and time, blood collection time, coordinates of harvest or distance and direction from nearest town, county of harvest, hunter-determined age class (juvenile, adult, or unsure) and sex (male, female, or unsure), any relevant comments, and hunter contact information. For organized hunts, samples were either stored at room temperature (feathers, Nobuto strips in 2018) or frozen to –18 C (heart samples and Nobuto strips in 2019 based on data indicating comparable test reliability; see also Dusek et al. 2014; Bevins et al. 2016) before submission. Otherwise, we provided hunters with pre-paid United Parcel Service shipping labels, freezer packs, and thermal-bubble mailers to keep samples cold during shipment for the next business day, until stored as above at –18 C. We confirmed age and sex from submitted feathers (DeStefano et al. 2014) before shipping samples to the University of Georgia (Athens, Georgia, USA) for analysis.

We also received grouse carcasses that were recently deceased for unknown reasons, were presumably sick based on abnormal behavior (e.g., unable to fly), or had reduced pectoral muscle and a prominent sternal keel. When possible, MNDNR staff obtained these carcasses and stored them frozen at –18 C until submission to the University of Minnesota, Veterinary Diagnostic Laboratory (St. Paul, Minnesota, USA). Pathologists certified by the American College of Veterinary Pathologists performed necropsies and looked for histologic lesions consistent with clinical WNV infection. Brain and heart samples from these carcasses were shipped to the Animal Health Diagnostic Center of Cornell University (Ithaca, New York, USA) to screen for WNV and eastern equine encephalitis by PCR.

Laboratory and data analysis

Nobuto strips were eluted per the manufacturer's instructions, yielding a dilution of 1:10. This dilution was approximate because of variable amounts of blood absorption on individual strips; however, all strips were treated similarly. Nobuto strip eluates were heat-inactivated at 56 C in 5% CO2 for 30 min, centrifuged at 12,000 × G for 4 min, and frozen at –20 C until testing. We tested eluates for anti-WNV antibodies with a plaque-reduction neutralization test (PRNT; Beaty et al. 1995), as described by Allison et al. (2004), except cultures were inactivated on d 5 after adsorption, rather than d 4, with 10%-buffered formalin and stained with 0.25–1.0% crystal violet for plaque visualization. The starting dilution was 1:20, and titers were expressed as the reciprocal of the highest eluate dilution that neutralized ≥90% WNV plaque-forming units (PRNT90) compared with control wells. We titrated samples with ≥90% neutralization against both WNV and St. Louis encephalitis virus (SLEV) to determine the causative virus (by a fourfold or greater PRNT90 titer); those that did not show at least a fourfold difference, presumably because of cross-reactivity (Nemeth et al. 2021) were considered antiflavivirus-antibody positive (Komar et al. 2001). Anti-flavivirus antibodies were interpreted as presumed antibodies to WNV for data analyses, based on the lack of documented SLEV circulation in the corresponding regions during the sampling years (CDC 2019).

To screen for circulating flaviviruses in heart tissue submitted to the Southeastern Cooperative Wildlife Disease Study (i.e., from hunter-harvested grouse), heart samples of approximately 0.5 cm3 were placed into 1 mL BA-1 medium, homogenized in a mixer mill (2 min for five cycles/s; Retsch MM 300, Haan, Germany), and centrifuged at 12,000 × G for 3 min. Homogenized heart sample (100 µL) was inoculated onto confluent, 2-d-old Vero cell-culture monolayers; incubated at 37 C in 5% CO2; and monitored daily for cytopathic effects for 10 d (Allison et al. 2004).

Viral RNA was extracted from all heart homogenates using the QiaAmp Viral RNA Mini Kit (Qiagen, Valencia, California, USA), per the manufacturer's protocol, and tested for WNV RNA by the VetMAX WNV TaqMan real-time reverse transcription (RT)-PCR kit (Applied Biosystems, Foster City, California, USA). We considered a cycle threshold value of ≤35 to be positive.

We used the number of antiflavivirus (i.e., anti-WNV) antibody-positive samples, relative to the total number of eluate samples tested, to separately calculate the apparent prevalence rate of WNV antibodies for each age class. We used the number of WNV real-time RT-PCR-positive heart samples, divided by the total number of heart samples tested by real-time RT-PCR, to calculate the active infection rate. Chi-square tests were performed in Microsoft Excel (Redmond, Washington, USA).

We mapped both seroprevalence and active infections using harvest location data (i.e., coordinates or approximate coordinates from hunter-submitted descriptions) in ArcMap version 10.6 (Redlands, California, USA). We used SaTScan version 9.6 (Boston, Massachusetts, USA; Kulldorff 2018) to look for clustering of positive test results in a space-time scan model. A Bernoulli cluster-scanning model using positive and negative results identified case clusters in which relative infection risk was greater than that in the surrounding area. We set maximum cluster size to 50% of the population at risk and set the window shape to circular.

In 2018, 117 hunters collected 273 samples from harvested Ruffed Grouse, with 213 collected within the 96.6-km sampling foci during 15 September–1 January (Fig. 1). Most returned kits contained all requested components, but 22 kits lacked hearts; 40 were missing some or all feathers for sex or age determination; and four lacked location information. In 2019, 166 hunters collected samples from 316 grouse harvested throughout the forest during 14 September–22 December (Fig. 1) and one fresh, road-killed grouse was collected 4 January 2020. Most returned kits were complete, but nine lacked hearts, 42 lacked feathers for age determination, 10 lacked feathers for sex determination, and five lacked location information. The median time to sample collection was 14 min, and 82.2% of 546 samples with reported collection times were collected ≤30 min after harvest.

Figure 1

The distribution within Minnesota of hunter-harvested Ruffed Grouse (Bonasa umbellus) samples and carcasses of presumably sick grouse submitted for testing during 2018 and 2019. WNV=West Nile virus.

Figure 1

The distribution within Minnesota of hunter-harvested Ruffed Grouse (Bonasa umbellus) samples and carcasses of presumably sick grouse submitted for testing during 2018 and 2019. WNV=West Nile virus.

Close modal

We detected antibodies to WNV or flavivirus in 12.5% (34/273) and 12.3% (39/317) of samples in 2018 and 2019, respectively. Anti-WNV antibodies were detected in 10 and three samples in 2018 and 2019, respectively. Antibodies to WNV were not distinguishable from those to SLEV in 24 and 36 samples in 2018 and 2019, respectively and, thus, were considered positive for anti-flavivirus antibodies. No heart samples were WNV-positive by real-time RT-PCR in either year.

Fourteen whole-grouse carcasses were submitted for necropsy because birds had been behaving oddly (seven), found dead (five), or were emaciated (two); all were negative for WNV. Two of these birds were included among the 317 hunter-submitted samples in 2019. Eleven were also tested for eastern equine encephalitis, and four were positive (Anderson et al. 2021).

Cohort composition and WNV prevalence

In 2018, we corrected hunter-determined age 50/211 times (24%) and sex 15/245 times (6%), excluding cases in which hunters were unsure or when feathers were not provided for verification. In 2019, we corrected hunter-determined age 53/186 times (28%) and sex 24/229 times (10%). In samples for which sex and age could both be verified, the male:female ratio was 53:47 in 2018 (n=212) and 64:36 (n=235) in 2019. Similarly, the juvenile:adult ratio of the verified sample in 2018 was 64:36 and 71:29 in 2019. We used these verified samples to examine WNV seroprevalence among cohorts.

We determined flavivirus (probably WNV) seroprevalence among verified cohorts for both years combined because sample sizes were small for some cohorts (range, 21–102). Seroprevalence in adults was similar to that in juveniles (15.1% vs. 11.6%, χ2=1.05, P=0.31), and males were similar to females (14.9% vs. 9.7%, χ2=2.59, P=0.11). Antibody prevalence in adult females (10.7%, n=56) was similar to that of juvenile females (10.1%, n=129), juvenile males (13.3%, n=172), and adult males (17.8%, n=90; χ2=3.08, P=0.38). However, we had low statistical power to detect differences: 650 individuals per cohort are needed to detect a 5% difference.

Spatial patterns

We found no statistically significant spatial patterns among all samples. However, we found a marginally significant cluster of positive cases about 20 km northeast of Hibbing, Minnesota (log-likelihood ratio, 8.44; P=0.07).

A low, but consistent, proportion of hunter-harvested Ruffed Grouse had antibodies to a flavivirus, probably WNV, each year, indicating that some birds previously infected with WNV survived to the fall. These data do not reveal whether birds were asymptomatic or experienced disease and recovered and do not indicate whether some died from infection before the fall. Species (populations) that are largely resistant to WNV have more-similar seroprevalence and infection rates than do those susceptible to disease-related mortality, so for WNV-susceptible species, inferences about infection rates are suspect without data on mortality (Walker et al. 2007). However, virus was not isolated and viral RNA was not detected via PCR from any submitted hearts, indicating birds were not actively infected when harvested. Furthermore, during our study, no grouse that were observed to be sick or dying from suspect causes and were submitted for testing had WNV. Thus, this study lacks evidence that Minnesota Ruffed Grouse developed WNV-associated morbidity and mortality, although elsewhere in the Great Lakes region, WNV-associated heart pathology has been reported in Ruffed Grouse (MIDNR 2017).

Interpretation of antibody prevalence can also be complicated by antibody-response duration (Gibbs et al. 2005). However, based on studies in other species, we would expect antibodies from summer infections of Ruffed Grouse to persist throughout the fall harvest season. For example, in seven raptor species, antibodies to WNV were protective for multiple years (Nemeth et al. 2008). Neutralizing antibody titers were detected for >36 mo in House Sparrows, with antibodies detectable in adults 1 mo after inoculation (Nemeth et al. 2009a). In Rock Pigeons (Columba livia), WNV-antibodies persisted ≥15 mo (Gibbs et al. 2005); and House Finches (Haemorhous mexicanus) had detectable antibodies for ≥28 wk (reviewed in Pérez-Ramírez et al. 2014). Thus, species may vary in the duration of antibody persistence and protection from reinfection (Gibbs et al. 2005).

The period of WNV detection in tissues and serum can vary among species, sample type, age cohorts, and detection methods. In House Sparrows experimentally inoculated with WNV, infectious virus persisted ≤43 d in tissues (Nemeth et al. 2009b). Virus was detected in most, but not all, skin, spleen, and kidney samples at 30 d postinoculation (DPI), but lesions were not histologically observed in juveniles at 30 or 65 DPI. Wheeler et al. (2012) detected viral RNA in House Sparrow sera for 2–7 wk, in the kidney up to 18 wk after infection, and in the spleen 12 wk after infection. Similarly, WNV RNA was detected in blood or organs >6 wk after infection in 34% of passerines and columbids examined (Reisen et al. 2006). However, in laboratory-inoculated Ruffed Grouse, virus was detected only in the kidney at 14 DPI (Nemeth et al. 2017). Lack of virus detection in hearts probably indicates that infections were cleared before our fall sampling. However, WNV may have been detectable in tissues that we did not sample.

Bird species with the highest WNV viremia titers usually, but not always, have the highest mortality rates (Pérez-Ramírez et al. 2014). “Super spreaders” develop high viremia titers without succumbing to infection; this occurs in several bird species (Pérez-Ramírez et al. 2014), which may amplify WNV transmission in nature. Among Galliformes, numerous species have low viremia titers; however, Red-legged Partridge (Alectoris rufa) and Greater Sage-grouse develop high viremia. Nevertheless, Greater Sage-grouse can survive infection; WNV-attributable mortality was estimated as 2.4–13.3%, with maximum possible rates of 8.2–28.9% in a wild population (Walker et al. 2007). Similarly, experimentally inoculated Ruffed Grouse developed high mean peak viremia titers (107 plaque-forming units/mL; Nemeth et al. 2017). We did not detect virus in the hearts of harvested birds and found no evidence of wild Ruffed Grouse sick or dying from WNV in our study. Summer sampling of wild birds would be better for virus detection but would present considerable challenges in obtaining adequate sample sizes, due to an inability to rely on hunters to collect samples, combined with low carcass recovery for testing because diseased birds may be predated and/or scavenged.

Cohort comparisons

We had low power to detect statistical differences among cohorts, with <650 birds/ cohort, and other studies have failed to find statistical differences among cohorts with similar sample sizes (Nemeth et al. 2021). However, in our study, adult males had 1.66× the seroprevalence of females. Seroprevalence differences between these cohorts might result from varied use of summer habitats, with corresponding differences in mosquito populations. In the summer in Maine, US, males use areas with higher woody stem density and lower Rubus (bramble) ground coverage than females with broods (Mangelinckx et al. 2018). It is unknown how Ruffed Grouse summer habitat use and mosquito populations vary in Minnesota or how those factors might affect WNV exposure.

Additionally, WNV seroprevalence might differ between adult Ruffed Grouse cohorts if female reproductive costs leave them in poorer condition to face immune challenges and other stressors than males, which do not care for eggs or broods. Corticosterone, a stress hormone in birds, is immunosuppressive, and WNV mortality increased in birds with corticosterone implants (Owen et al. 2012). Lower survival of female Ruffed Grouse with broods (69%) than those without broods and males (98%) has been reported in Maine (Mangelinckx et al. 2018), which is consistent with male-skewed sex ratios among harvested Ruffed Grouse in our study and others (Dorney 1963; Davis and Stoll 1973). Although sex differences in the survival of WNV infection have not been observed in the highly susceptible American Crow (Yaremych et al. 2004), both parents care for young.

Adult female Ruffed Grouse had similar WNV-antibody prevalence to that of juveniles. This finding is inconsistent with higher WNV-associated mortality rates in juveniles (Austin et al. 2004; Sovada et al. 2008; Cox et al. 2015), which would be expected to correspond with lower seroprevalence in (surviving) juveniles. We might expect exposure of females and their broods to be similar over the summer because they use the same areas, are in close contact, and thus, probably have similar exposure to WNV-infected mosquitoes. Bird-to-bird transmission has also been reported in some (McLean et al. 2001; Langevin et al. 2001; Komar et al. 2003), but not all, species examined in experimental settings (Nemeth et al. 2009a; Pérez-Ramírez et al. 2014). Alternatively, chicks might have similar antibody prevalence to their mothers because in some species, antibodies can be passed maternally through eggs (Gibbs et al. 2005; see review in Ahlers and Goodman 2018). However, antibodies detectable in chicks did not last >42 d in any species evaluated (Perez-Ramirez et al. 2014), and Ruffed Grouse hatching in the Great Lakes region is expected to peak in mid-June (Larson et al. 2003), approximately 90 d before our sampling began. Similar antibody prevalence between juveniles, who could only recently be exposed to WNV, and adult females, might indicate that antibody titers are not maintained for life in adult Ruffed Grouse as they are in some other species (Gibbs et al. 2005; Nemeth et al. 2009b; Pérez-Ramírez et al. 2014). Additional research on the persistence of WNV antibodies is necessary to improve understanding of adult titers over time.

Comparisons with other regions

Prevalence of antibodies consistent with WNV in Minnesota Ruffed Grouse varied less than that found in Pennsylvania, in which statewide serosurveys from hunter-harvested birds during 2015–17 indicated apparent prevalence rates of 13.3% (n=188), 22.6% (n=230), and 2.8% (n=145), each year, respectively (Nemeth et al. 2021). We cannot know, from the available data, whether differences in seroprevalence were due to differences in infection rates and/or differences in survival rates of infected Ruffed Grouse. Either way, WNV does not appear to cause additive mortality in Minnesota Ruffed Grouse. The Ruffed Grouse population in Minnesota is cycling around a stable long-term average (Roy 2017), whereas grouse numbers have been declining for decades in Pennsylvania (Stauffer et al. 2018). Furthermore, Minnesota has abundant forested habitat in a variety of successional stages that support high grouse densities (Zimmerman et al. 2009; Kouffeld et al. 2013), compared with Pennsylvania in which poor-quality habitat occurs where WNV prevalence is greater (Stauffer et al. 2018).

Viruses and other pathogens become a concern for wildlife populations when a population is naïve and newly exposed or when other stressors reduce resistance to immune challenges. Interactions between emerging diseases and land-use can produce complex effects on survival of wild birds (George et al. 2015). In Pennsylvania, habitat scarcity might be a stressor for Ruffed Grouse that could reduce resistance to WNV (Stauffer et al. 2018). Providing abundant, high-quality habitat for Ruffed Grouse currently appears to be our best option for protecting populations from WNV because birds in good condition have stronger immune responses. Other options, such as reducing mosquito populations at a broad scale, vaccinating Ruffed Grouse (Nemeth et al. 2017) at meaningful levels, and eliminating viruses, are not currently feasible. However, by providing the best habitat possible, we will produce grouse more likely to be robust to a variety of stressors, including viruses.

We appreciate the efforts of all the hunters that submitted samples, without which, this study would not be possible. We are grateful to the Pineridge Grouse Camp, Bowen Lodge and Northwoods Bird Dogs, the Akeley Grouse Hunt, the Ruffed Grouse Society, and contributing staff and students of Itasca Community College; DNR staff and retirees; as well as friends and family, with a special thanks to Charlie Tucker and Matt Breuer for their large sample contributions, as well as Earl Johnson and Jerry Havel for assistance at the Pineridge Grouse Camp. Erik Hildebrand, Patrick Hagen, Margaret Dexter, and Lindsey Peterson assisted with sample kit building. Christopher Jennelle performed spatial analysis. This study was funded, in part, by the Ruffed Grouse Society, in addition to funding from Federal Aid in Wildlife Restoration. We are also thankful for the thoughtful reviews provided by three anonymous reviewers.

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